3005
Journal of Cell Science 107, 3005-3013 (1994)
Printed in Great Britain © The Company of Biologists Limited 1994
Meiosis-specific cell cycle regulation in maturing Xenopus oocytes
Keita Ohsumi*, Wako Sawada and Takeo Kishimoto
Laboratory of Cell and Developmental Biology, Faculty of Biosciences, Tokyo Institute of Technology, 4259 Nagatsuta, Midori-ku,
Yokohama 227, Japan
*Author for correspondence
SUMMARY
Meiotic cell cycles differ from mitotic cell cycles in that the
former lack S-phase in the interphase between meiosis I
and meiosis II. To obtain clues for mechanisms involved in
the cell cycle regulation unique to meiosis, we have
examined changes in chromosomal morphology and H1
kinase activity during a meiotic period from metaphase I
(MI) to metaphase II (MII) in Xenopus oocytes. Using populations of oocytes that underwent germinal vesicle
breakdown (GVBD) within a 10 minute interval, we found
that the kinase activity declined gradually during the 60
minute period after GVBD and then increased steadily
during the following 80 minute interval, showing remarkable differences from the rapid drop and biphasic increase
of the kinase activity in intermitotic periods (Solomon et al.
(1990) Cell 63, 1013-1024; Dasso and Newport (1990) Cell
61, 811-823). We also found that the exit from MI lagged,
by more than 30 minutes, behind the time of lowest H1
kinase activity, whereas the two events took place concomitantly at the end of meiosis II and mitosis. Conse-
quently, the H1 kinase activity was already increasing
during the first meiotic division. When H1 kinase activation at MII was delayed by a transient inhibition of protein
synthesis after GVBD, oocytes were able to support
formation of interphase nuclei and DNA replication
between the first meiotic division and the MII arrest, indicating that the cell cycle entered S-phase between meiosis
I and meiosis II. These results strongly suggest that the
machinery required for entering S-phase has been established in maturing oocytes by the end of meiosis I. The lack
of S-phase in oocyte meiotic interphase, therefore, should
be ascribed to cell cycle regulation that enables the transition from meiosis I to meiosis II without S-phase. The asynchrony between the inactivation of H1 kinase activity and
the completion of meiosis I may be involved in the regulation of this unique feature of the meiotic cell cycle.
INTRODUCTION
meiosis II proceeds without entering S-phase, the regulation of
cdc2 kinase in meiotic cycles may differ from that in mitotic
cycles. Although the mechanisms for coupling of the cdc2
kinase activation with the completion of S-phase have been
investigated (for review see Murray, 1992; Enoch and Nurse,
1990; Kumagai and Dunphy, 1991; Smythe and Newport,
1992), kinase activation uncoupled from S-phase has not been
studied.
Apart from the regulation of cdc2 kinase, the unique absence
of S-phase between the two successive M-phases in meiotic
cycles could be explained by inability of oocytes to replicate
chromosomal DNA, since, in Xenopus laevis, immature, fully
grown oocytes are incompetent for replicating chromosomal or
double-stranded DNA, whereas mature eggs are highly
competent (Gurdon, 1967; Benbow and Ford, 1975; Cox and
Leno, 1990). Although the ability to replicate chromosomal
DNA appears in maturing oocytes after germinal vesicle
breakdown (GVBD) (Gurdon, 1967), the precise timing of its
occurrence has not yet been determined. Interestingly,
maturing mouse oocytes treated with puromycin at meiotic
metaphase I (MI) undergo nucleus formation but not DNA
replication after meiosis I, and further protein synthesis is
required for this nucleus to undergo DNA replication (Clarke
Maturation-promoting factor (MPF) was first found in the
oocytes of frogs (Masui and Markert, 1971) and starfish
(Kishimoto and Kanatani, 1976) as the cytoplasmic activity
that can induce the resumption of meiotic divisions arrested at
the G2/M border. Since the demonstration in frogs (Wasserman
and Smith, 1978; Gerhart et al., 1984) and starfish (Kishimoto
et al., 1982) that MPF activity also appears in cleaving
embryos concomitantly with mitosis, it has been postulated
that both meiosis and mitosis (M-phase) are driven by the same
cytoplasmic activity, MPF. Recent progress in the study of the
regulation of M-phase has substantiated that MPF is attributable to p34cdc2 in a complex with cyclin B (cyclin B-dependent
cdc2 kinase; cdc2 kinase) (for review, see Murray and
Kirschner, 1989b; Nurse, 1990). The protein kinase activity of
the complex is routinely quantified using histone H1 as a
substrate (Lohka et al., 1988; Labbé et al., 1989; Gautier et al.,
1989, 1990). The similarities rather than the differences
between meiosis and mitosis have been emphasized by the
demonstration that both M-phases are induced as a result of
activation of cdc2 kinase. However, considering that mitosis
requires the completion of S-phase, whereas the transition to
Key words: meiosis, cell cycle regulation, cdc2 kinase, DNA
replication, Xenopus oocyte
3006 K. Ohsumi, W. Sawada and T. Kishimoto
and Masui, 1983; Clarke et al., 1988; Hashimoto and
Kishimoto, 1988). Thus it is possible that the activity to
replicate chromosomal DNA may not have been established in
maturing Xenopus oocytes at the time they enter meiotic interphase or interkinesis.
For a comparison of the regulation of meiotic cycles to that
of mitotic cycles, Xenopus oocytes provide a system as useful
as oocytes of marine invertebrates (Westendorf et al., 1989;
Hunt et al., 1992). The advantages of the Xenopus system
include the ability to induce the resumption of meiotic
divisions in vitro by hormonal stimulation (for review see
Masui and Clarke, 1979) and the accumulation of extensive
information on the regulatory mechanisms of mitotic cycles,
which has been obtained by the use of cell-free systems with
egg extracts (Lohka and Maller, 1985; Miake-Lye and
Kirschner, 1985; Murray and Kirschner, 1989a, Murray et al.,
1989). However, in Xenopus, information on the meiotic cycles
after GVBD is limited, in part because GVBD occurs asynchronously even in oocytes from a single female. Thus the
description of the time course of meiotic progression from
GVBD to meiotic metaphase II (MII) was not obtained until
the study by Gerhart et al. (1984), who synchronized maturing
oocytes by selecting populations of oocytes that reached
GVBD within short intervals.
To know the regulatory mechanisms for the cell cycle transition to meiosis II without passing through S-phase, we have
extended the work by Gerhart et al. (1984) by examining
detailed changes of H1 kinase activity and p34cdc2/cyclin B
complex along with meiotic progression from MI to MII. The
results demonstrate that the regulation of cdc2 kinase in the
post-GVBD period differs from that in mitosis, and that the
exit from MI considerably lags behind the preceding kinase
inactivation and consequently occurs when kinase activation
for meiosis II is already under way. The delayed exit from MI
in the presence of active cdc2 kinase may be required to
prevent meiotic cycles of maturing oocytes from entering Sphase, since delay in activation of kinase activity can lead to
S-phase after meiosis I.
MATERIALS AND METHODS
Oocytes, eggs, embryos and sperm
Mature females and males of Xenopus laevis were purchased from the
Nippon Bio-supp. Center, Tokyo. Female frogs were injected with 50
units of pregnant mare serum gonadotropin 3 to 7 days before ovary
dissection or induction of ovulation. Excised ovary fragments were
treated with MMR (100 mM NaCl, 2 mM KCl, 1 mM MgCl2, 2 mM
CaCl2, 0.1 mM EDTA, 5 mM HEPES, pH 7.8; Newport and
Kirschner, 1982) containing 0.2% collagenase (Type S-1, Nitta
Gelatin Corp., Osaka) and 10 µg/ml progesterone for 2 hours at room
temperature with continuous agitation. After extensive washing with
MMR, fully-grown oocytes at stage VI (Dumont, 1972) were
collected and incubated at 22°C. Populations of synchronized oocytes
were obtained by collecting oocytes on which the spot had appeared
within 10 minute intervals (Gerhart et al., 1984) and incubated at
22°C. In some experiments, maturing oocytes were transferred to
MMR containing 10 µg/ml cycloheximide or 50 µg/ml aphidicolin at
30 minutes post-GVBD. Some oocytes received an injection of 50 nl
of sperm suspension (5×104 sperm/µl sucrose solution; 250 mM
sucrose, 5 mM MgCl 2, 10 mM Tris-HCl, pH 7.4) containing 200 µM
biotin-14-dATP (Gibco, BRL) with or without 1 or 5 mg/ml
puromycin at 25-35 minutes post-GVBD.
Eggs and embryos were obtained and dejellied as described previously (Ohsumi and Katagiri, 1991; Ohsumi et al., 1993). Briefly,
female frogs were induced to ovulate by injection of human chorionic
gonadotropin (300-500 units/female), and eggs were artificially
inseminated. Eggs and embryos were dejellied with 2.5% thioglycolic
acid (pH 8.2). In some experiments, dejellied eggs were activated by
stimulation with an electrical shock (60 mA dc, 1 second, 2 times).
Cleaving embryos were synchronized by collecting embryos on which
the tips of the first cleavage furrow had reached the equatorial region
within 5 minutes intervals.
Sperm were demembranated with 0.05% lysolecithin according to
Lohka and Masui (1983) and stored at −80°C.
Microscopy
The nucleus and chromosomes of oocytes and eggs were observed
according to the method of Gerhart et al. (1984). Oocytes and eggs
were fixed with 3% formalin in MMR for at least 1 hour and their
white spot region was dissected with watchmaker’s forceps in 50%
glycerol (MMR) containing 10 µg/ml diamidinophenylindole (DAPI).
The patch was mounted on a glass slide with a coverslip so that the
plasma membrane side faces up. Sperm nuclei injected into oocytes
and embryonic nuclei were examined by squashing the oocytes and
embryos as described previously (Ohsumi et al., 1986).
To detect the incorporation of biotin-dATP (Blow and Watson,
1987), oocytes were squashed under a coverslip with a minimum
amount of MMR, and immediately frozen in liquid nitrogen. After the
coverslip was removed, frozen samples were fixed with Carnoy’s
fixative for 3 minutes, rinsed twice with ethanol and air-dried. The
preparations were rinsed three times in PBS and incubated with 1/200diluted Texas Red-conjugated streptavidin (Amersham Corp.) for 30
minutes at room temperature, followed by staining with 10 µg/ml
DAPI.
These preparations were observed under an epifluorescence microscope (Zeiss Axioscope).
Histone H1 kinase assay
An individual oocyte, egg and embryo was homogenized with 50 µl
of an extraction buffer (EB; 100 mM KCl, 15 mM MgCl2, 5 mM
EGTA, 80 mM β-glycerophosphate, pH 7.4, 1 mM phenylmethylsulfonyl fluoride, 100 µg/ml leupeptin, 100 µg/ml aprotinin) and immediately frozen in liquid nitrogen. The homogenates were centrifuged
(10,000 g, 3 minutes, 2°C) and 40 µl of supernatant was recovered
for each sample. The extracts were mixed with 5 µl of an ATP solution
(80 mM β-glycerophosphate, pH 7.4, 15 mM MgCl2, 4 mM ATP)
containing 8 µg histone H1 (Boehringer Mannheim) and 1 µCi [γ32P]ATP, and incubated for 30 minutes at 22°C. Reactions were
stopped by the addition of SDS-sample buffer and boiling for 2
minutes. Samples were run on 15% polyacrylamide gels, and stained
with Coomassie Blue, dried, and autoradiographed on X-ray film. For
measurement of 32P incorporation, each H1 band was excised from
Coomassie Blue-stained gels and counted by the Cerenkov method.
Immunoblotting
Ten oocytes were homogenized with 100 µl of EB and their cytoplasmic extracts were prepared as for H1 kinase assay. The extracts
were incubated with 10 µl of p13suc1-beads (5 µg of p13suc1/µl of gel;
Ookata et al., 1992) for 30 minutes at 4°C with continuous gentle
agitation. After washing with EB, the beads were mixed with 15 µl
of SDS-sample buffer, and boiled for 2 minutes. The samples were
run on 12.5% polyacrylamide gels and blotted onto an Immobilon
membrane (Millipore) according to the method of Towbin et al.
(1979). After blocking with 3% skimmed milk, the membrane was
incubated with primary antibodies for 1 hour at room temperature.
Antibodies used in this study were anti-PSTAIR (gift from Drs M.
Yamashita and Y. Nagahama), anti-phosphotyrosine (gift from Dr G.
Peaucellier), anti-Xenopus cyclins B1 and B2 (gift from Drs T. Hunt
and J. L. Maller), and anti-Xenopus cdc2 and cdk2 (gift from Dr T.
Meiotic cell cycles in Xenopus oocytes 3007
Hunt). The membrane was then incubated with alkaline phosphataseconjugated secondary antibodies for 1 hour at room temperature.
Signals were visualized by a BCIP/NBT phosphatase substrate system
(KPL).
RESULTS
The meiotic cell cycle after GVBD of Xenopus
oocytes is highly synchronous
Under our experimental conditions, a white spot in the animal
pole region, an external indicator of GVBD, appeared asynchronously in oocytes at 2.5 to 5 hours after the hormonal
administration. Populations of maturing oocytes that had
undergone GVBD within a 10 minute interval were collected
and incubated at 22°C. These oocytes were fixed at various
times after GVBD for observation of chromosomal morphology according to the method of Gerhart et al. (1984). Changes
of oocyte chromosomes during the post-GVBD period are
presented in Fig. 1. Condensed chromosomes, which had been
scattering in the animal pole region at the time of GVBD,
gathered to be aligned in circular arrangement by 60 minutes
post-GVBD (Fig. 1A). This state persisted until 90 minutes
post-GVBD, at which time chromosomes were divided into
two groups (Fig. 1B). In the following 10 minutes, chromosomes of each group aggregated to form a chromatin mass; the
chromosomes of the first polar body were distinguishable from
those remaining in the oocyte as the former was encapsulated
by a membrane (Fig. 1C). By 130 minutes post-GVBD,
discrete chromosomes in a circular arrangement became
visible again (Fig. 1D), and this state was maintained for at
least 3 hours. This sequential change of oocyte chromosomes
during the post-GVBD period is essentially the same as that
observed by Gerhart et al. (1984). Although information on the
meiotic spindle is not provided in these observations, the
change of chromosomal morphology presented in Fig. 1 reasonably represents the meiotic progression from MI to MII, i.e.
MI (Fig. 1A), first meiotic anaphase (Fig. 1B), interkinesis or
meiotic interphase (Fig. 1C), and MII (Fig. 1D). During the
period from first meiotic anaphase to MII, the chromosomes
remaining in an oocyte never formed an interphase nucleus
(see also Gard, 1992); the formation of an interphase nucleus,
had one formed, would have been discernible as decondensed
chromatin with a smooth contour due to nuclear membrane
formation (cf. Fig. 4A).
Based on the change of chromosomal morphology, the transition through meiosis after GVBD is plotted in Fig. 1E. In this
figure, MI and MII are put together as metaphase, since they
were not always distinguishable, and results from 7 batches of
oocytes from 6 females are summarized. The steep decrease
and increase in the ratio of metaphase oocytes indicate that
both the exit from MI and attainment of MII can occur synchronously after GVBD among oocytes from different females.
It may be noted that the percentage of oocytes in interkinesis
does not reach 100% at any time point. This, however, does
not mean an arrest of meiosis at MI in a subpopulation of
oocytes, since histological observation of paraffin sections
showed that all oocytes at 150 minutes post-GVBD had the
polar body near the M-phase nucleus (K. Ohsumi, unpublished
result). Thus the failure to observe all oocytes in interkinesis
at one time should be ascribed to the shortness of the interki-
Fig. 1. The time course of meiosis from MI to MII in Xenopus
oocytes. (A-D) Maturing oocytes were fixed at various times after
GVBD and stained with DAPI for observation of chromosomal
morphology according to the method of Gerhart et al. (1984).
Chromosomes of oocytes fixed at 60 (A), 90 (B), 100 (C), and 130
minutes (D) post-GVBD represent MI, first meiotic anaphase,
meiotic interphase or interkinesis, and MII, respectively. In (B) and
(C) two chromosome masses, which were observed at different
depths from the oocyte surface are presented in combination. Bar, 10
µm. (E) Based on the chromosomal morphology, the ratios of MI and
MII (s), interkinesis (j), and anaphase oocytes (n) are plotted as a
function of time. Results from seven batches of oocytes from six
females are summarized up. The numbers at the top represent the
number of oocytes observed.
nesis period. The duration of interkinesis including first
meiotic telophase was estimated to be approximately 20
minutes from the interval between the time points at which
50% of oocytes leave MI and reach MII (Fig. 1E). These results
clearly demonstrate that in Xenopus oocytes the progression
through meiosis after GVBD occurs highly synchronously
under our defined conditions.
Oscillation of H1 kinase activity during the MI/MII
period
To know the precise time course of the oscillation of MPF
activity during the meiotic period from MI to MII, oocytes
were individually examined for histone H1 kinase activity at
various times after GVBD. Fig. 2A plots kinetics of the kinase
activity during the post-GVBD period as a function of time.
3008 K. Ohsumi, W. Sawada and T. Kishimoto
(Solomon et al., 1990), was absent from immature oocytes
before progesterone treatment, and its amount changed in proportion to the extent of H1 kinase activity during the postGVBD period. Similarly, the amount of both cyclin B1 and the
upper band of cyclin B2 was proportional to the kinase activity,
although it was reproducibly found that the amount of cyclin
B1 reaches a minimum slightly earlier than the amount of
cyclin B2. Thus, the oscillation of H1 kinase activity during
the MI/MII period is obviously correlated with the state of
p34cdc2/cyclin B complex. Another point notable in Fig. 2B is
that phosphotyrosyl p34cdc2 is not detected during the MI/MII
period, whereas it is found in immature oocytes and the mitotic
interphase of fertilized eggs (Ferrell et al., 1991; W. Sawada,
unpublished result).
Fig. 2. The changes in H1 kinase activity and p34cdc2/cyclin B
complex during the post-GVBD period from MI to MII in Xenopus
oocytes. (A) The kinetics of H1 kinase activity of maturing oocytes
as a function of time. Maturing oocytes were individually examined
for the kinase activity at various times after GVBD as described in
Materials and Methods. Three oocytes were examined at each time
point. IM and AC indicate immature oocytes before the hormonal
stimulation and activated eggs at 15 minutes after the activation,
respectively. Each point represents mean ± s.d. (B) Immunoblotting
of p34cdc2/cyclin B complex from oocytes at various times after
GVBD. Ooplasmic extracts were prepared from ten oocytes, and
p34cdc2/cyclin B complex was isolated by the use of p13suc1-beads
for immunoblotting with anti-PSTAIR, anti-phosphotyrosine, antiXenopus cyclin B1, and anti-Xenopus cyclin B2 antibodies as
described in Materials and Methods.
H1 kinase activity, which had been at a high level at GVBD,
decreased after GVBD, and reached a minimum level, comparable to that of activated eggs, at 80 minutes post-GVBD. Then
the activity increased again to an MII level during the
following 80 minutes, and the high level was maintained for
at least 3 hours. This time course of the oscillation of H1 kinase
activity during the post-GVBD period roughly corresponds to
that of the oscillation of MPF activity (Gerhart et al., 1984).
To confirm that the oscillation of H1 kinase activity is attributed to the regulation of cdc2 kinase, we examined changes in
p34cdc2/cyclin B complex during the post-GVBD period.
p34cdc2/cyclin B complex was isolated for immunoblotting
from ooplasmic extracts by using p13sucl-beads. As shown in
Fig. 2B, five differently migrating bands on an SDS-PAGE gel
were recognized with anti-PSTAIR antibody. Of these, the
upper three and lower two bands were regarded as p34cdc2 and
p32cdk2, respectively, because of their reactivity to anti-cdc2
and anti-cdk2 antibodies (data not shown). The lowest band of
p34cdc2, which has been thought to be the active form
The exit from MI lags behind the H1 kinase
inactivation
The comparison of the timing of the exit from MI (Fig. 1E)
and the inactivation of H1 kinase following GVBD (Fig. 2A)
suggests that there is a considerable time lag between the two
events. To ascertain the lag, a population of synchronously
maturing oocytes was divided into two groups, one for observation of chromosomal morphology and the other for examination of H1 kinase activity. The results from oocytes of the
same female are presented in Fig. 3A. This figure clearly shows
that the exit from MI lags more than 30 minutes behind the
time at which H1 kinase activity decreases to minimum, and
consequently occurs when H1 kinase activity has already
started to re-increase.
For comparison, the temporal relationship between the exit
from metaphase and H1 kinase inactivation was examined for
meiosis II and mitosis in the same way. The results show that
the two events occur almost concomitantly in meiosis II and
mitosis (Fig. 3B,C). Thus the delayed exit from metaphase is
unique to meiosis I.
Kinetics of H1 kinase activity in the MI/MII period
differs from that in mitosis
The detailed change in H1 kinase activity presented in Fig. 3A
also reveals that the kinetics of H1 kinase activity in the
MI/MII period is distinct from that in intermitotic periods in
both activation and inactivation of the kinase. In the MI/MII
period, the kinase activity declines gradually for more than 60
minutes and subsequently increases steadily, corresponding to
the gradual decrease and increase of the cyclin B amount. In
mitosis, the kinase activity drops abruptly within 15 minutes
owing to rapid degradation of cyclin (Fig. 3C; Murray and
Kirschner, 1989a) and increases in a biphasic manner while
cyclin accumulates continuously (Solomon et al., 1990; Dasso
and Newport, 1990).
Oocytes inhibited from the transition to MII enter Sphase
The finding that the first meiotic division occurs in the presence
of H1 kinase activity prompted us to examine effects on
meiosis of blocking the reactivation of MPF at MII by protein
synthesis inhibition (Gerhart et al., 1984). When maturing
oocytes were incubated with cycloheximide (10 µg/ml) at 30
minutes post-GVBD, the H1 kinase inactivation was not
affected but its reactivation was completely blocked (Fig. 3D).
In these oocytes, the first meiotic division occurred on schedule
Meiotic cell cycles in Xenopus oocytes 3009
Fig. 3. The temporal relationship between H1 kinase inactivation and
the exit from metaphase in meiosis I and II, and in mitosis. The
changes in H1 kinase activity (d) and the metaphase indexes (s) in
meiosis I (A,D), meiosis II (B), and the 2nd mitosis (C) were
examined in a batch of oocytes, eggs and embryos from a single
female. Populations of oocytes maturing synchronously in the
absence (A) or presence (D) of 10 µg/ml cycloheximide, activated
eggs (B), cleaving embryos (C) were divided into two groups, and
both H1 kinase activity and chromosomal morphology were
examined at the same times after GVBD (A,D), the activation by
electric shock (B), and the first cleavage furrow formation (C) as
described in Materials and Methods. For each time point, H1 kinase
activity and chromosomal morphology were examined on 3 to 5 and
more than 10 samples, respectively. Each point of H1 kinase activity
represents mean ± s.d. Arrow in (D) indicates the time of the
addition of cycloheximide.
(Fig. 3D), but subsequently the oocyte chromosomes formed a
nucleus (Fig. 4A) instead of remaining at the interkinesis state
(Fig. 1C). The same result was obtained by injecting
puromycin (250 ng/oocyte) into maturing oocytes at 30
minutes post-GVBD (data not shown).
We then examined whether or not the nuclei formed in the
puromycin-treated oocytes undergoes DNA replication. For
this study, we used demembranated sperm nuclei, which are
known to undergo nuclear transformation as well as DNA
replication in concordance with the resident nuclei when
injected into oocytes and eggs (Graham, 1966; Moriya and
Katagiri, 1976). Maturing oocytes were injected with sperm
nuclei, biotin-dATP, and puromycin (250 ng/oocyte) at 30
minutes post-GVBD. Injected oocytes were squashed at 150
minutes post-GVBD for both observation of sperm nuclear
morphology and examination of their incorporation of biotindATP by the use of Texas Red-streptavidin. The results
presented in Fig. 4 confirm that sperm nuclei injected into
oocytes are transformed to a morphology similar to that of the
resident oocyte nucleus: formation at 150 minutes post-GVBD
of interphase nuclei (Fig. 4B,D) and metaphase chromosomes
(Fig. 4F) in puromycin-injected and control oocytes, respectively. The results also show an obvious incorporation of
biotin-dATP by the nuclei formed in puromycin-injected
oocytes (Fig. 4C). This incorporation was blocked with aphidicolin (Fig. 4E), suggesting that it was due to the replication but
not the repair synthesis of DNA. The occurrence of DNA
synthesis on the resident oocyte nucleus under these conditions
was confirmed by examining oocytes injected with puromycin
and biotin-dATP alone (data not shown). In the protein
synthesis inhibition experiments, we found that in oocytes
injected with a lower dose of puromycin (50 ng/oocyte) at 30
minutes post-GVBD, protein synthesis was inhibited by more
than 95% at 60 minutes post-GVBD, but was restored by 50%
at 150 minutes post-GVBD. It was shown by immunoblotting
that in these oocytes both cyclin B1 and B2 disappeared by 90
minutes post-GVBD but reappeared by 180 minutes postGVBD, accompanying H1 kinase activation, and at 120
minutes post-GVBD, p34cdc2 and p32cdk2 were in essentially
the same forms as those at mitotic interphase, which included
the phosphotyrosyl p34cdc2 (W. Sawada, unpublished result).
When sperm nuclei were injected into these oocytes in which
protein synthesis was transiently inhibited, they transformed
into interphase nuclei by 120 minutes post-GVBD, and then
into condensed metaphase chromosomes by 180 minutes postGVBD (Fig. 4H). The incorporation of biotin-dATP by these
chromosomes (Fig. 4I) indicates that they have passed S-phase.
Interestingly, the replicated sperm-derived chromosomes (Fig.
4H) are highly condensed, distinct from non-replicated spermderived chromosomes (Fig. 4F). The metaphase state was
maintained for at least three hours presumably by the action of
the cytostatic factor, indicating that the metaphase is delayed
MII. We conclude from these results that maturing oocytes
enter S-phase after the exit from meiosis I when the entry into
meiosis II is blocked or delayed by protein synthesis inhibition.
DISCUSSION
We have shown in this study that maturing Xenopus oocytes
undergo nuclear formation and the replication of chromosomal
DNA after meiosis I, if the activation of H1 kinase at MII is
blocked or delayed by protein synthesis inhibition. Since both
nuclear formation and DNA replication is induced by the inhibition of protein synthesis after GVBD, machinery required for
entry into S-phase must be already established in maturing
oocytes by the end of meiosis I; the ability to replicate chromosomal DNA seems to appear in maturing oocytes around
the time of GVBD and depends on newly synthesized proteins,
since oocytes injected with puromycin 30 minutes before
GVBD subsequently underwent nuclear formation but not
DNA replication (K. Ohsumi et al., unpublished result). These
data strongly indicate that the absence of S-phase in the interkinesis of maturing oocytes is not simply due to incompetence
of the oocytes for entering S-phase, but should be ascribed to
a meiosis-specific cell cycle regulation that enables the transition from meiosis I to meiosis II while bypassing S-phase.
cdc2 kinase regulation unique to the MI/MII period
Entry into and exit from mitosis are shown to require the acti-
3010 K. Ohsumi, W. Sawada and T. Kishimoto
Fig. 4. The formation of interphase nuclei and occurrence of DNA
replication in maturing oocytes blocked from the transition to MII by
protein synthesis inhibition. Maturing oocytes were incubated with
10 µg/ml cycloheximide (A) or injected with demembranated sperm
and biotin-dATP together with puromycin at the dose of 250 (B-E), 0
(F,G), or 50 ng (H,I) per oocyte at 30 minutes post-GVBD. Some
oocytes were incubated in the presence of 50 µg/ml aphidicolin
(D,E). Oocytes were squashed and fixed on glass slides at 150 (A-G)
or 180 minutes (H,I) post-GVBD, and stained with DAPI
(A,B,D,F,H) and Texas Red-streptavidin (C,E,G,I) for examination
of chromosomal morphology and incorporation of biotin-dATP by
the chromosomes. The oocyte nucleus transforms into an interphase
nucleus after the exit from MI in oocytes inhibited from protein
synthesis after GVBD by cycloheximide (A). Similarly, sperm nuclei
transform into interphase nuclei (B,D) accompanying incorporation
of biotin-dATP (C), which is inhibited by aphidicolin (E), in oocytes
injected with the high dose of puromycin (250 ng/oocyte), whereas
they transform into metaphase chromosomes in control oocytes (F)
without the incorporation of biotin-dATP (G). In oocytes injected
with the low dose of puromycin (50 ng/oocyte), sperm nuclei
transform into highly condensed metaphase chromosomes (H) via
interphase nuclear formation accompanying biotin-dATP
incorporation (I). Bar, 10 µm.
vation and inactivation of the kinase activity of p34cdc2, respectively (Murray and Kirschner, 1989a; Murray et al., 1989;
Minshull et al., 1989). We have confirmed that in maturing
oocytes H1 kinase activity decreases during the meiotic period
from MI to MII (Labbé et al., 1988; Kanki and Donoghue,
1991; Yew et al., 1992; Izumi et al., 1992), consistent with the
decline in MPF activity (Gerhart et al., 1984; Daar et al., 1991).
Furthermore, our detailed analysis revealed that the kinetics of
H1 kinase activity in the MI/MII period is markedly different
from that in the intermitotic periods of activated eggs and
cleaving embryos, as discussed below.
The inactivation of cdc2 kinase, which is caused by cyclin
degradation (Murray et al., 1989; Draetta et al., 1989), occurs
rapidly within 15 minutes at the end of meiosis II and mitosis
(Murray et al., 1989; this study). We showed that in the MI/MII
period H1 kinase activity decreases more slowly, over a period
of more than 60 minutes. Although H1 kinase activity was
examined with the whole oocyte cytoplasm in this study, most
of the activity can be attributed to p34cdc2/cyclin B complex
for the following reasons: more than 90% of the kinase activity
was absorbed with p13suc1-beads (W. Sawada, unpublished
result), and the contribution of p34cdc2 and p32cdk2 complexed
with cyclin A seems negligible because the amount of cyclin
A is small during this period (Kobayashi et al., 1991). We also
showed that the decrease in the amounts of cyclin B1 and B2
is consistent with the decline of H1 kinase activity. This was
confirmed when H1 kinase activity was measured on each of
cyclin B1 and B2 complexes that were obtained by immunoprecipitation with specific antibodies (K. Ohsumi, unpublished
result). We suggest therefore that the slower inactivation of
cdc2 kinase in the MI/MII period is attributable to the longer
lasting, gradual decrease of the cyclin B amount. The slower
decrease in the amount of cyclin B is thought to be due to lower
rates of its degradation, because both synthetic rates and steady
state levels of cyclin B do not differ appreciably between
maturing oocytes at GVBD and mature eggs at MII (Kobayashi
et al., 1991). This implies that mechanisms involved in cyclin
degradation may be different between the MI/MII period and
mitosis. It is also noteworthy that in the MI/MII period the
amounts of cyclin B1 and B2 reach their minima at different
times; the turning point from a decrease to an increase of
protein levels comes earlier for cyclin B1 than for cyclin B2.
This difference is possibly related to the difference between
Meiotic cell cycles in Xenopus oocytes 3011
cyclin B1 and B2 in their requirement of binding to p34cdc2 for
degradation (Stewart et al., 1994).
The activation of p34cdc2 requires posttranslational modifications in addition to its association with cyclin B. In particular, a phosphotyrosine, which renders the kinase inactivated,
needs to be removed from p34cdc2 by cdc25 phosphatase
(Kumagai and Dunphy, 1991; Izumi et al., 1992; Hoffmann et
al., 1993). This dephosphorylation is thought to be a ratelimiting step in the activation of the kinase after the cyclin
binding. Consequently, while the accumulation of cyclin B is
continuous, the activation of cdc2 kinase occurs abruptly,
being delayed after the concentration of cyclin B reaches a
critical threshold (Solomon et al., 1990). Thus, the increase in
H1 kinase activity at the transition to mitosis appears to be
biphasic, the initial slow rise followed by the rapid increase
(Solomon et al., 1990; Dasso and Newport, 1990). We found,
however, that in the MI/MII period H1 kinase activity
increases monotonously in parallel with the continuous
increase in the amount of p34cdc2-associated cyclin B. We also
found that the phosphotyrosine-containing form of p34cdc2
does not occur during the MI/MII period (see also Ferrell et
al., 1991). This finding is consistent with the observation that
cdc25 phosphatase remains in its highly phosphorylated, active
form throughout the MI/MII period (Izumi et al., 1992).
Together these results suggest that the inactivation by tyrosine
phosphorylation is not involved in the regulation of cdc2
kinase during the MI/MII period, thereby the subsequent activation of the kinase for meiosis II would be facilitated. The
swift activation of cdc2 kinase in the MI/MII period is thought
to be required for the transition to meiosis II without passing
through S-phase because its delay was shown to induce Sphase after meiosis I.
The activation of cdc2 kinase for meiosis II is also distinct
from that for mitosis in that the former requires the synthesis
of proteins other than cyclin B including at least mos protein
(Daar et al., 1991; Kanki and Donoghue, 1991; Yew et al.,
1992), the active component of cytostatic factor, which stabilizes MPF (Sagata et al., 1989). Very recently, after the submission of this manuscript, Furuno et al. (1994) have demonstrated that mos-mediated activation of cdc2 kinase is required
for preventing maturing Xenopus oocytes from entering Sphase after meiosis I. In contrast, it has been reported that new
synthesis of cyclin B is not required for the meiotic transition
to MII; the fraction of cyclin B that survives meiosis I is sufficient for the transition (Minshull et al., 1991). Consistent with
these observations, we have found that considerable amounts
of both cyclin B1 and B2 remain even after H1 kinase inactivation in the MI/MII period, whereas virtually all of the cyclins
disappear at the end of meiosis II (see Fig. 2B).
Asynchronized progression of meiosis with H1
kinase oscillation in the MI/MII period
Another striking finding in this study is that in the MI/MII
period, the meiotic progression is obviously not synchronized
with the oscillation of H1 kinase activity: the exit from MI lags
considerably behind the inactivation of H1 kinase and takes
place when the increase in kinase activity required for meiosis
II has already started. This asynchrony cannot be ascribed to
the large size of amphibian oocytes, because in eggs and
embryos of the same size, the exit from MII and mitotic
metaphase were shown to occur concomitantly with H1 kinase
inactivation, and besides, a similar asynchrony seems to occur
in maturing oocytes of an European starfish (see Fig. 2 in Galas
et al., 1994). Rather, taking into account that the MI lasts far
beyond the H1 kinase inactivation (see Fig. 3A), it seems very
likely that the metaphase state of chromosomes is maintained
against the decline of cdc2 kinase activity in meiosis I. It is
noteworthy in this regard that cyclin B associates with tubulin
and is partly localized on the metaphase spindle and chromosomes in meiosis I of maturing starfish oocytes (Ookata et al.,
1992, 1993) and that mos protein has been shown to colocalize with tubulin in mitotic spindles (Zhou et al., 1991). Thus,
it seems possible that the p34cdc2 localized on the MI spindle
together with mos kinase might remain active even after cytoplasmic p34cdc2 is inactivated, and could be involved in the
maintenance of MI. In support of this notion, the treatment of
maturing oocytes after GVBD with either potent kinase
inhibitors (6-DMAP and staurosporine) or anti-tubulin drugs
(vinblastin and colcemid) caused the condensed chromosomes
at MI to transform into an aggregate of chromosomes, an
interkinesis-like state, concomitantly with the H1 kinase inactivation, suggesting that both the activities of kinases and
microtubules are required to maintain the MI state of oocyte
chromosomes (K. Ohsumi, unpublished result).
In spite of the asynchrony in the oscillation of the cytoplasmic p34cdc2 kinase activity, the exit from MI was shown to
occur highly synchronously at a certain time after GVBD. This
indicates that the anaphase transition in meiosis I is strictly
controlled and must be causally related to the oscillation of
cytoplasmic cdc2 kinase, although their relationship would be
more indirect and complicated than postulated in mitosis.
Although the mechanism that triggers the anaphase transition
in meiosis I is unknown at present, the meiotic division in the
presence of H1 kinase activity is not inconceivable since it has
been demonstrated that anaphase transition can be induced separately from cdc2 kinase inactivation under experimental conditions (Holloway et al., 1993; Surana et al., 1993).
In conclusion, the meiotic cycle from MI to MII is characterized by the specific regulation of p34cdc2 and the asynchrony
of meiotic progression and the kinase oscillation. Both of these
characteristics of meiosis-specific regulation may be involved
in mechanisms that ensure that the transition from meiosis I to
meiosis II occurs without oocytes entering S-phase.
We thank Drs T. Hunt (ICRF), J. L. Maller (University of
Colorado), G. Peaucellier (Roscoff), M. Yamashita and Y. Nagahama
(National Institute for Basic Biology) for generous gifts of antibodies;
Dr M. J. Lohka (Uniersity of Calgary) for critical reading of the manuscript; and S. Hisanaga and K. Tachibana for helpful discussions.
This work was supported by grants from the Ministry of Education,
Science and Culture of Japan and the Asahi Glass Foundation to T.K.
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(Received 10 May 1994 - Accepted 15 July 1994)